active pin1 is a key target of all-trans retinoic acid in...
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Targeted therapy has changed cancer treatment, but blocking a single pathway is often ineffective against solid tumors, especially aggressive or drug-resistant ones, because of activation of redundant and/or alter-native oncogenic pathways1. Thus, knowing how to block the multiple cancer-driving pathways simultaneously remains a major challenge. A common and central signaling mechanism in oncogenic pathways is proline-directed phosphorylation (pSer/Thr-Pro)2. Numerous oncogenes and tumor suppressors are either directly regulated by (Supplementary Fig. 1) and/or trigger signal pathways involving such phosphorylation2,3. Notably, the same kinases often phosphorylate both oncogenes and tumor suppressors to control their function. The prolyl isomerase (PPIase) Pin1 has a critical role in coordinating these multiple phosphorylation events to oncogenesis2,3.
Proline uniquely adopts cis and trans conformations, and its isomer-ization is catalyzed by PPIases4, including the unique PPIase Pin1 (refs. 2,5,6). Using its WW domain, Pin1 binds to specific pSer/Thr-Pro motif(s), in which its PPIase domain catalyzes cis-trans isomerization of certain pSer/Thr-Pro motifs5, which can be detected by cis- and trans-specific antibodies6. Pin1 is commonly overexpressed and/or activated in human cancers, which correlates with poor outcomes3,7. In contrast, Pin1 polymorphisms that lower Pin1 expression are asso-ciated with reduced cancer risk compared to the normal population8.
Moreover, Pin1 deficiency in mice prevents tumorigenesis, even that induced by activated oncogenes such as Erbb2 (encoding HER2) or Kras (encoding Ras)9, whereas Pin1 overexpression disrupts cell cycle coordination and leads to centrosome amplification, chromo-some instability, and cancer development in cell and animal models of breast cancer10. Pin1 activates at least 32 oncogenes and growth-promoting proteins, and inactivates at least 19 tumor suppressors and growth-inhibiting proteins2,3,11–20 (Supplementary Fig. 1). Thus, Pin1 can amplify oncogenic pathways by simultaneously activating oncogenes and inactivating tumor suppressors. Pin1 also has a fundamental role in driving expansion and tumorigenesis of cancer stem cells21–23, a major source of cancer resistance1. These studies suggest that Pin1 inhibitors could have the unique and desirable ability to block multiple cancer-driving pathways and inhibit cancer stem cells at the same time2,3,24, especially given that Pin1-knockout (KO) mice develop normally without obvious defects for an extended period of time25,26.
However, the available Pin1 inhibitors either lack the required specifi-city and/or potency or cannot efficiently enter cells to inhibit Pin1 func-tion in vivo3,27. Here we developed mechanism-based high-throughput screening for compounds targeting active Pin1. We found that ATRA (tretinoin) directly and selectively binds, inhibits and ultimately
1Division of Translational Therapeutics, Department of Medicine, Beth Israel Deaconess Medical Center (BIDMC), Harvard Medical School, Boston, Massachusetts, USA. 2Department of Medicine, BIDMC, Harvard Medical School, Boston, Massachusetts, USA. 3Cancer Research Institute, Beth Israel Deaconess Cancer Center, Harvard Medical School, Boston, Massachusetts, USA. 4Department of Pathology, BIDMC, Harvard Medical School, Boston, Massachusetts, USA. 5Department of Molecular Biosciences, University of Texas, Austin, Texas, USA. 6Division of Gerontology, Department of Medicine, BIDMC, Harvard Medical School, Boston, Massachusetts, USA. 7Department of Biological Chemistry and Molecular Pharmacology, Harvard Medical School, Boston, Massachusetts, USA. 8Department of Systems Biology, Harvard Medical School, Boston, Massachusetts, USA. 9Department of Internal Medicine, University of São Paulo, Ribeirão Preto, Brazil. 10Department of Biomedicine and Prevention, Tor Vergata University and Santa Lucia Foundation, Rome, Italy. 11Present address: Department of Natural Products and Experimental Therapeutics, University of Hawaii Cancer Center, Honolulu, Hawaii, USA. Correspondence should be addressed to K.P.L. ([email protected]) or X.Z.Z. ([email protected]).
Received 5 December 2014; accepted 16 March 2015; published online 13 April 2015; doi:10.1038/nm.3839
Active Pin1 is a key target of all-trans retinoic acid in acute promyelocytic leukemia and breast cancerShuo Wei1–3, Shingo Kozono1–3, Lev Kats2–4, Morris Nechama1–3, Wenzong Li5, Jlenia Guarnerio2–4, Manli Luo1–3, Mi-Hyeon You6, Yandan Yao1–3, Asami Kondo1–3, Hai Hu2,3, Gunes Bozkurt7, Nathan J Moerke8, Shugeng Cao7,11, Markus Reschke2–4, Chun-Hau Chen1–3, Eduardo M Rego9, Francesco Lo-Coco10, Lewis C Cantley2,3,8, Tae Ho Lee6, Hao Wu7, Yan Zhang5, Pier Paolo Pandolfi2–4, Xiao Zhen Zhou1–3 & Kun Ping Lu1–3
A common key regulator of oncogenic signaling pathways in multiple tumor types is the unique isomerase Pin1. However, available Pin1 inhibitors lack the required specificity and potency for inhibiting Pin1 function in vivo. By using mechanism-based screening, here we find that all-trans retinoic acid (ATRA)—a therapy for acute promyelocytic leukemia (APL) that is considered the first example of targeted therapy in cancer, but whose drug target remains elusive—inhibits and degrades active Pin1 selectively in cancer cells by directly binding to the substrate phosphate- and proline-binding pockets in the Pin1 active site. ATRA-induced Pin1 ablation degrades the protein encoded by the fusion oncogene PML–RARA and treats APL in APL cell and animal models as well as in human patients. ATRA-induced Pin1 ablation also potently inhibits triple-negative breast cancer cell growth in human cells and in animal models by acting on many Pin1 substrate oncogenes and tumor suppressors. Thus, ATRA simultaneously blocks multiple Pin1-regulated cancer-driving pathways, an attractive property for treating aggressive and drug-resistant tumors.
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degrades active Pin1, thereby exerting potent anticancer activity against APL and triple-negative breast cancer (TNBC) by simultane-ously blocking multiple Pin1-regulated cancer-driving pathways.
RESULTSMechanism-based screening for Pin1 inhibitorsPhosphorylation of Pin1 at S71 by the tumor suppressor DAPK1 (ref. 28) inhibits Pin1 catalytic activity and oncogenic function by blocking a phosphorylated substrate from entering the active site7 (Supplementary Fig. 2a). Such phosphorylation would probably also prevent Pin1 from binding to pTide, a high-affinity, substrate- mimicking peptide inhibitor (Bth-d-phos.Thr-Pip-Nal; Kd = 1.2 nM) that cannot enter cells7,29 (Supplementary Fig. 2b). Indeed, fluorescently labeled pTide bound to Pin1, but not to FK506- binding protein 12 (FKBP12), and to the Pin1 PPIase domain, but not to its WW domain (Supplementary Fig. 2c–f). pTide also bound to the nonphosphorylatable Pin1 S71A mutant, but not to its phospho- mimicking S71E mutant; binding depended on Pin1 active site resi-dues that mediate phosphate binding, such as K63 and R69, and those that mediate Pro recognition, such as L122, M130, Q131, and F134 (ref. 29) (Supplementary Fig. 2g). Thus, we developed and performed a fluorescence polarization–based high-throughput screen (FP-HTS) for chemical compounds that could compete with pTide for binding to nonphosphorylated (and thus active) Pin1. Out of ~8,200 com-pounds screened, 13-cis-retinoic acid (13cRA) was the top hit on the basis of the lowest z-score (Fig. 1a).
13cRA and its isomer, ATRA (Fig. 1b,c), bound to Pin1 in the FP assay, with ATRA being more potent than 13cRA after a short period of incu-bation (Ki = 1.16 and 0.58 µM, respectively, calculated using an equation as previously described30) (Supplementary Fig. 3a), but this differ-ence disappeared after a longer incubation (Supplementary Fig. 3b,c). These results suggest that Pin1 may mainly bind to the trans form (ATRA) but that it can bind to the cis form (13cRA) after it is con-verted to the trans form, which does occur over time both in vitro and in vivo31. The ATRA–Pin1 interaction was confirmed using a differ-ent fluorescence-labeled pTide probe (Supplementary Fig. 2h). As [3H]ATRA has been used as a photoaffinity-labeling reagent to cova-lently and specifically tag ATRA-binding proteins32, we performed photoaffinity labeling of Pin1 with [3H]ATRA, and subsequently detected the binding using SDS-containing gels. [3H]ATRA directly bound to Pin1 (Kd = 0.80 µM) (Fig. 1d,e and Supplementary Fig. 3a). Moreover, ATRA and 13cRA fully inhibited the PPIase activity of Pin1, with Ki values of 0.82 and 2.37 µM, respectively, which were similar to those from the FP assay (Fig. 1f and Supplementary Fig. 3a,d,e), but neither compound inhibited cyclophilin and FKBP12, other major PPIases (Supplementary Fig. 3f,g). Thus, ATRA is a submicromolar Pin1 inhibitor.
ATRA binds to the Pin1 active siteTo determine whether the carboxylic acid in ATRA serves as an alter-native to the phosphate group for binding to Pin1, several structurally similar retinoids with substituted carboxylic or aromatic groups and
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Figure 1 Mechanism-based screening identifies ATRA as a submicromolar Pin1 inhibitor that binds to the Pin1 active site. (a) Summary plot of FP-HTS for Pin1 inhibitors, with 13-cis-retinoic acid having the lowest z-score, as determined by folds of s.d. below the mean of each screening plate. (b,c) Structures of cis (13cRA)- (b) and trans (ATRA)-retinoic acid (c). (d,e) [3H]ATRA binds to Pin1 in a dose-dependent manner. (d) Pin1 incubated with various concentrations of [3H]ATRA followed by UV exposure before SDS-gel and radiography. (e) Quantified Pin1-bound [3H]ATRA signals, as plotted against ATRA concentrations. Mean ± s.d. of three experiments. (f) Inhibition of Pin1 catalytic activity by ATRA or 13cRA, as measured by PPIase assay. Mean ± s.d. of two experiments. (g) FP readout of Pin1 incubated with different concentrations of various compounds for 0.5 h. pTide-HiLyte Fluor 488 was added to Pin1, followed by incubation of different concentrations of compounds indicated for 0.5 h, before FP readout (mean ± s.d. of three experiments). (h) After ATRA soaking, strong electron density was observed at the Pin1 active site in the co-crystal. (i) In the ATRA-Pin1 co-crystal structure, ATRA-Pin1 binding (middle) is mediated by salt bridges between the carboxylic acid of ATRA and K63 and R69 residues (right), as well as by hydrophobic interaction between aromatic moiety of ATRA and L122, M130, Q131 and F134 residues (left). The PDB code for the Pin1-ATRA structure is 4TNS.
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the new generations of retinoids, fenretinide33 and bexarotene34, were tested for Pin1 binding. ATRA was the most potent against Pin1 out of those tested (Fig. 1g and Supplementary Fig. 4a). Notably, car-boxylic acid group (-COOH)-substituted retinoids, including retinol (-OH), retinyl acetate (-OCOCH3) and retinal (-CHO) were totally inactive (Fig. 1g and Supplementary Fig. 4a). In line with this, fen-retinide and bexarotene showed only marginal Pin1 binding (Fig. 1g and Supplementary Fig. 4a), which might be due to the lack of a carboxyl group and/or modifications to target retinoic acid receptors (RARs), retinoid X receptors (RXRs), or others33–35.
To understand how ATRA inhibits Pin1 catalytic activity, we deter-mined the co-crystal structure of ATRA and the Pin1 PPIase domain
(Supplementary Fig. 4b and Supplementary Table 1). After ATRA soaking, strong electron density was observed at the Pin1 active site (Fig. 1h). The most-well-defined region of ATRA was its carboxyl group, which formed salt bridges with Pin1’s critical catalytic resi-dues K63 and R69 (Fig. 1i), both of which are essential for binding the phosphate group in the Pin1 substrate29. At the high resolution of 1.3 Å, two alternative conformations of R69 were visible, both of which were within the distance range needed for salt bridge forma-tion with the carboxyl group of ATRA. The trimethylcyclohexene ring of ATRA was sandwiched within Pin1’s hydrophobic Pro-binding pocket, which is formed by L122, M130, Q131 and F134 (ref. 29) (Fig. 1i). Notably, the binding modes of ATRA and pTide overlapped
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Figure 2 ATRA causes Pin1 degradation and inhibits its oncogenic function in cells. (a) Cell count assay of WT and Pin1-KO MEFs (left) or Pin1-KO MEFs reconstituted with either WT or W34/K63A Pin1 (right) treated with the indicated concentrations of ATRA for 72 h. (b) Immunoblot of WT and Pin1-KO MEFs (top) or Pin1-KO MEFs reconstituted with WT or W34/K63A Pin1 (bottom) treated with the indicated concentrations of 13cRA or ATRA for 72 h. β-actin, loading control. (c) Top, gel of Pin1 mRNA levels, as detected by RT-PCR, in MEFs treated with the indicated concentrations of ATRA or 13cRA for 72 h (top). Bottom, graph showing quantification of the data. GADPH, control. (d) Top, immunoblot of Pin1 abundance in MEFs treated with the indicated concentrations of ATRA for 48 h followed by co-treatment of ATRA and either 10 µM MG132 or DMSO (control) for an additional 24 h. Bottom, graph showing quantification of the data. (e) Top, immunoblot of Pin1 abundance from MEFs treated with ATRA, 13cRA (both 50 µM), or DMSO for 24 h followed by cycloheximide (CHX) chase for the indicated times. Bottom, graph showing quantification of the data. (f) Images of immunostained NIH 3T3 cells stably expressing Flag-tagged Pin1 or empty vector (Ctrl) and treated with the indicated concentrations of ATRA for 72 h. Red arrows, centrosomes; scale bars, 10 µm. (g) Quantification of cells containing over two centrosomes, from three experiments with over 100 cells in each. (h) Reporter gene assay of SKBR3 cells co-transfected with a cyclin D1 promoter–driven luciferase construct and either Flag-Pin1 or Ctrl and treated with the indicated concentrations of ATRA for 72 h. (i,j) Picture (i) and foci quantification (n = 3) (j) of SKBR3 cells co-transfected with Flag-Pin1 or empty vector (Ctrl) and treated with the indicated concentrations of ATRA for 48 h. Foci formation assay was applied. *P < 0.05, **P < 0.01, ***P < 0.001, as determined by Student’s t-test. Error bars, means ± s.d. Results are representative of three experiments unless stated otherwise.
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(Fig. 1i and Supplementary Fig. 2b). Thus, by mimicking the pSer/Thr-Pro motif in a substrate, the carboxylic and aromatic moie-ties of ATRA bind to the substrate phosphate- and proline-binding pockets of the Pin1 active site, respectively. These structural require-ments were also consistent with our findings that the carboxyl group of ATRA was required for binding to Pin1, and that fenretinide and bexarotene were less potent than ATRA in binding Pin1 (Fig. 1g).
ATRA induces Pin1 degradation and inhibits its functionTo determine whether ATRA inhibits Pin1 activity in cells, we first compared its effects on the proliferation of Pin1-KO (Pin1−/−) and wild-type (WT, Pin1+/+) mouse embryonic fibroblasts (MEFs). Relatively high concentrations of ATRA were required to inhibit the growth of Pin1 WT MEFs, and Pin1-KO cells were more resistant than Pin1 WT MEFs to ATRA (Fig. 2a). Susceptibility to ATRA was fully restored by re-expressing Pin1 but not its inactive W34/K63A mutant (Fig. 2a). Notably, ATRA also dose-dependently downregulated levels of WT but not mutant Pin1 protein (Fig. 2b). ATRA had no obvious effects on Pin1 mRNA levels (Fig. 2c). ATRA reduced protein levels of both exog-enous and endogenous Pin1, but not that of the W34/K63A mutant, which did not bind ATRA (Fig. 2b and Supplementary Fig. 2g). ATRA-induced Pin1 degradation was suppressed by the proteasome inhibitor MG132 (Fig. 2d). Both ATRA and 13cRA reduced Pin1 protein half-life (Fig. 2e), but ATRA was more potent (Fig. 2b,e).
Next we examined the effects of ATRA on well-documented onco-genic phenotypes induced by Pin1 overexpression, such as centro-some amplification10, activation of the cyclin D1 promoter11 and
enhanced foci formation; all of these phenotypes are inhibited by DAPK1-mediated Pin1 phosphorylation at S71 (ref. 7). ATRA dose-dependently and fully inhibited the ability of Pin1 overexpression to induce centrosome amplification in NIH 3T3 MEFs (Fig. 2f,g), as well as its ability to activate the cyclin D1 promoter and enhance foci formation in SKBR3 cells (Fig. 2h–j). Thus, ATRA induces Pin1 degradation and inhibits its oncogenic function.
Pin1 is a key target of ATRA in APL cellsATRA is approved to treat APL, in which it activates RARs to induce APL cell differentiation and also causes degradation of the fusion protein promyelocytic leukemia–retinoic acid receptor α (PML–RAR-α) to inhibit self-renewal of APL stem cells36–38. However, ATRA-induced RAR-α activation can be decoupled from its ability to degrade PML–RAR-α and treat APL39,40. Notably, retinoid analogs that potently activate RARs and induce leukemia cell differentiation, but fail to induce PML–RAR-α degradation, also fail to inhibit self-renewal of leukemia stem cells and treat APL40. Moreover, ATRA’s ability to activate RARs cannot readily explain its ability to destabilize other oncogenic molecules, including cyclin D1 (ref. 41) and NF-κB42, or its ability to stabilize tumor-suppressive molecules such as Smad43. Thus, the cellular target(s) of ATRA that mediate its anticancer effects remain elusive.
To examine the role of RARs in ATRA-directed degradation of PML–RAR-α, we used a pan-RAR agonist, AC-93253, and a pan-RAR
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Figure 3 Pin1 is a critical target for ATRA to induce PML–RAR-α degradation and inhibit proliferation in APL cells. (a,b) NB4 cells were stably infected by lentivirus expressing shPin1 and WT or W34/K63A Flag-Pin1, followed by immunoblotting for PML–RAR-α or Pin1 (a) or counting cell number over time (b) (mean ± s.d. of three experiments). (c–e) NB4 cells stably infected by lentivirus expressing shPin1 and WT or W34/K63A Flag-Pin1 were subjected to the CHX chase, followed by immunoblotting for PML–RAR-α and Pin1 (c,d), with quantification shown in e (mean ± s.d. of three experiments). (f) Hierarchical cluster of the differential expression profiles showed similar profiles in ATRA-treated and Pin1-KD NB4 cells. NB4 cells were treated with ATRA or DMSO for 72 h, or subjected to doxycycline (shPin1)- or mock (vec)-induced shPin1 KD for 72 h, followed by mRNA extraction and microarray analysis. Colors, fold changes. (g–i) Immunodeficient NSG mice were transplanted with 5 × 105 human APL NB4 cells stably carrying inducible Tet-on shPin1 and, 5 d later, they received doxycycline-containing food to induce Pin1 KD; this was followed by examining PML–RAR-α and Pin1 in the bone marrow (g), measuring spleen size (h) (mean ± s.d. of four mice) and evaluating disease-free survival (i) of transplanted mice. (j) Bone marrow samples from the mice labeled with A, B and C were subjected to immunoblotting for PML–RAR-α and Pin1.**P < 0.01, as determined by Student’s t-test.
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inhibitor, Ro-415253, both of which are struc-turally distinct from ATRA (Supplementary Fig. 5a) and exhibit the expected ability to activate or inhibit transcription of RAR down-stream targets, respectively (Supplementary Fig. 5b). Ro-415253 showed minimal Pin1 binding and AC-93253 showed no binding (Supplementary Fig. 5c). Ro-415253 neither prevented ATRA from inducing degradation of Pin1 or PML–RAR-α (Supplementary Fig. 5d) nor inhibited the growth of NB4 human APL cells44 (Supplementary Fig. 5e), whereas AC-93253 did not mimic ATRA-induced Pin1 or PML–RAR-α degradation (Supplementary Fig. 5f). These RAR-independent ATRA effects were also confirmed using RAR-α, RAR-β, and RAR-γ triple-KO MEFs40, in which ATRA induced degradation of PML–RAR-α and Pin1 similarly to what was observed in WT controls (Supplementary Fig. 5g,h).
ATRA-induced PML–RAR-α degradation is associated with phos-phorylation of the Ser581-Pro motif of PML–RAR-α (ref. 39), which corresponds to the Pin1 binding site pSer77-Pro in RAR-α (ref. 45). Because Pin1 binds to and increases the protein stability of numer-ous oncogenes2,3 (Supplementary Fig. 1), we hypothesized that Pin1 might bind to the pS581-Pro motif in PML–RAR-α and stabilize it, thereby promoting APL cell growth. Indeed, Pin1 interacted with PML–RAR-α, and the S581A but not the S578A mutation abolished this interaction (Supplementary Fig. 6a) and reduced PML–RAR-α stability2,3 (Supplementary Fig. 6b,c). Moreover, Pin1 knockdown
(KD) using a validated shRNA-containing lentivirus7 reduced PML–RAR-α stability and inhibited APL cell growth compared to pLKO vector control; both effects were rescued by re-expression of shRNA-resistant Pin1, but not of its inactive mutant (Fig. 3a–e). In contrast to PML–RAR-α, Pin1 interacted much less with promyelo-cytic leukemia zinc finger ortholog (PLZF)–RAR-α (Supplementary Fig. 7a), and Pin1 KD only marginally reduced the protein stability of PLZF–RAR-α, as compared with PML–RAR-α (Supplementary Fig. 7b–e). Although future experiments are needed to define the underlying mechanisms, these results are consistent with the fact that APL induced by PLZF–RAR-α is usually resistant to ATRA36–38.
To expand our investigation beyond PML–RAR-α, we compared genome-wide gene expression profiles of ATRA- or DMSO-treated NB4 cells and NB4 cells stably expressing either pLKO vector control or Pin1 shRNA using microarrays that covered coding and noncoding transcripts in the human whole genome. Clustering analysis revealed a similarity between ATRA-treated and Pin1 shRNA-expressing cells. 528 genes were differentially expressed both in Pin1-KD cells and ATRA-treated cells, as compared with their respective controls. 304 genes were upregulated, including many growth suppressors (for example,
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Figure 4 Inhibition of Pin1 by ATRA or other compounds causes PML–RAR-α degradation and treats APL in cell and mouse models and human subjects. (a,b) NB4 cells were treated with ATRA, various Pin1 inhibitors, RAR inhibitor or RAR activator for 72 h, followed by immunoblotting for PML–RAR-α and Pin1 (a) or Giemsa staining (b, top) or FACS with CD14 and CD11b (b, bottom) for detecting APL cell differentiation. Scale bar, 10 µm. (c–e) APL cell differentiation status as measured by Giemsa staining (c, top) or FACS with Gr-1 and Mac-1 (c, bottom); PML–RAR-α and Pin1 expression in the bone marrow (d); and spleen size (e) of sublethally irradiated C57BL/6J mice (n = 10) transplanted with 1 × 106 APL cells isolated from CTSG-PML-RARA transgenic mice and, 5 d later, treated with ATRA-releasing implants, EGCG, Juglone or placebo for 3 weeks. Mac-1 is the same as CD11b; scale bar, 10 µm. (f–h) Bone marrow samples from healthy controls (n = 24) or APL patients before (n = 19) or after the treatment with ATRA for 3 (n = 3) or 10 days (n = 3) or APL patients in complete remission (n = 17) were immunostained with Pin1- and PML-specific antibodies (f). Red arrows, PML–RAR-α/PML diffusely distributed to the entire nucleus in APL cells containing more Pin1; yellow arrows, PML–RAR-α/PML localized to the PML body in APL cells containing less Pin1. Scale bars, 5 µm. (g,h) Relative expression of Pin1 in the nucleus (g) and PML–RAR-α in the nuclear plasma outside of the PML nuclear body (h) were semi-quantified (mean ± s.d.). *P < 0.05, **P < 0.01, ***P < 0.001, as determined by Student’s t-test.
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PDCD4 and SORL1) and 224 were downregulated, including many growth stimulators (for example, CCL2, SPP1, IL1B and IL8) in both Pin1-KD cells and ATRA-treated cells (Fig. 3f and Supplementary Table 2). Thus, both PML–RAR-α gene-specific and genome-wide analyses support the idea that ATRA inhibits Pin1 in APL cells.
We corroborated these in vitro results in animal studies using sublethally irradiated immunodeficient NOD scid gamma (NSG) mice transplanted with NB4 cells stably expressing an inducible tetracycline-controlled (Tet-on) shPin1 (ref. 46). When doxycycline-containing food was given 5 d after transplantation and throughout the remaining course of the experiment, Pin1 and PML–RAR-α expres-sion decreased in the bone marrow (Fig. 3g). In contrast to mice given control food, which exhibited splenomegaly, mice fed doxycycline displayed normal spleen size (Fig. 3h and Supplementary Fig. 8a). Doxycycline-fed mice also contained fewer human CD45-expressing NB4 cells in the bone marrow (Supplementary Fig. 8b–d). Disease-free survival of doxycycline-fed mice was also substantially extended compared to that of mice fed control chow (Fig. 3i). Notably, in one doxycycline-fed mouse that died early (i.e., before the other mice), Pin1 and PML–RAR-α were expressed in amounts close to those in mice not fed doxycycline (Fig. 3j), thereby supporting the role of Pin1 and its effects on PML–RAR-α in survival of mice with APL. Thus, as is the case with ATRA, inducible Pin1 KD alone is sufficient to cause PML–RAR-α degradation and treat APL in animal models.
ATRA and Pin1 inhibition suppress APL growthWe compared the effects of ATRA to the effects of three structurally distinct Pin1 inhibitors (PiB47, EGCG48 and Juglone49) on NB4 human APL cells in vitro. These three inhibitors are less potent than ATRA and also have other targets and toxicities27. Similarly to ATRA, these agents dose-dependently reduced PML–RAR-α expression in NB4 human APL cells, but they inhib-ited Pin1 without degrading it (Fig. 4a). However, in contrast to ATRA or the pan-RAR activator, neither these Pin1 inhibitors nor Pin1 shRNA induced NB4 human APL cell differentiation (Fig. 4b). These results were further supported by the observation that ATRA but not Pin1 shRNA induced the expression of RAR target genes (Supplementary Fig. 5i); the minimal effect of Pin1 shRNA could be attributed to the stabilization of the RAR protein mediated by Pin1 shRNA, as shown previously45.
To examine the effects of these Pin1 inhibitors on APL in vivo, we retro-orbitally injected sublethally irradiated B6 mice with APL cells isolated from CTSG–PML–RARA transgenic mice, which express human PML–RAR-α under the control of the myeloid/promyelocytic–specific cathepsin G gene promoter50, and then treated them 5 d later with EGCG (intraperitoneally (i.p.)) or Juglone (intravenously (i.v.)), or with ATRA-releasing pellets (5 mg over 21 d in one pellet implanted subcutaneously in the back of mice). 20 d after the start of treatment, we analyzed the differentiation of APL cells from bone
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Figure 5 ATRA ablates active Pin1 and thereby turns off oncogenes and turns on tumor suppressors in breast cancer. (a,b) Various human normal and breast cancer cells treated with ATRA for 72 h and assayed with colorimetric MTT assay (three experiments) (a) or directly subjected without the treatment to immunoprecipitation (IP)/immunoblotting for detecting Pin1 and its S71 phosphorylation (b). (c) The inverse correlation of Pin1 and DAPK1 in human TNBC tissues shown by immunohistochemical analysis (n = 48 TNBC tissues). Scale bars, 50 µm. (d–f) Immunoblotting of proteins expressed in various breast cells treated with different concentrations of ATRA for 72 h (d), various breast cells stably expressing tetracycline-inducible (Tet-on) Pin1 shRNA and treated with tetracycline for different times to induce Pin1 KD (e), and TNBC cells treated with tetracycline for different times after reconstitution of shRNA-resistant Pin1 or its W34/K63A mutant (f). Error bars, mean ± s.d.
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marrow by flow cytometry. ATRA, but neither EGCG nor Juglone, induced APL cell differentiation (Fig. 4c). Moreover, ATRA, but neither EGCG nor Juglone, reduced Pin1 levels in the bone marrow (Fig. 4d), although the reduction was not as profound as observed in vitro (Fig. 4a), probably because of the presence of normal cells in bone marrow, which were usually more resistant to ATRA (Figs. 2a and 5a,b). Nevertheless, all three Pin1 inhibitors effectively reduced PML–RAR-α protein expression in the bone marrow (Fig. 4d) and treated APL, with spleen weights nearly at basal levels (Fig. 4e and Supplementary Fig. 8e). Unlike ATRA-treated animals, EGCG- or Juglone-treated mice were rather sick, probably because of the fact that EGCG and Juglone have other toxic effects27. These results are consistent with the previous findings that ATRA’s ability to activate RARs and induce leukemia cell differentiation can be uncoupled from its ability to degrade PML–RAR-α and treat APL39,40.
Next we determined whether ATRA treatment degrades Pin1 and PML–RAR-α in APL cells in humans. We used double immuno-staining with antibodies against Pin1 and PML to detect Pin1 and
PML–RAR-α abundance and localization in cells from the bone marrow of healthy individuals, APL patients before or after ATRA treatment for 3 or 10 d, or individuals in complete APL remission (Supplementary Table 3), as described7,51. In contrast to healthy controls, Pin1 and PML–RAR-α were markedly overexpressed and distributed throughout the entire nuclei of cells from all subjects with APL who were examined before treatment (Fig. 4f–h). After ATRA treatment, PML–RAR-α levels were markedly reduced, with fluores-cence signal detected mainly in PML nuclear bodies (Fig. 4f). Of note, reduced fluorescence signal, mainly in PML nuclear bodies, repre-sents endogenous PML protein and reflects a good ATRA response51. Importantly, ATRA treatment caused a time-dependent reduction of Pin1 and PML–RAR-α expression to ~40% or <10% after 3 or 10 d of treatment, respectively (Fig. 4f–h). Notably, PML–RAR-α/PML stain-ing colocalized with Pin1 staining in APL cells. PML–RAR-α/PML was still diffusely distributed throughout the entire nuclei of APL cells containing more Pin1 (Fig. 4f), but it was almost exclusively localized to PML bodies (probably reflecting endogenous PML) in APL cells
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Figure 6 ATRA exerts potent anticancer activity against TNBC in vivo by ablating Pin1 and thereby blocking multiple cancer pathways simultaneously. (a–d) Shown are tumor sizes (a,c, top; measured weekly for 7 weeks), curves of tumor volume plotted over time (a,c, bottom) and representative immunoblots of Pin1 and cyclin D1 expression in xenograft tumors (b,d) from nude mice flank-inoculated with 2 × 106 MDA-MB-231 (a,b) or MDA-MB-468 cells (c,d) and implanted with ATRA-releasing (5 or 10 mg over 21 d) or placebo pellets 1 week later (arrows). n = 8 for a, 6 for c. (e) Tumor sizes (top; measured weekly for 4 weeks) and quantitative curves of tumor volume (bottom) from nude mice flank-inoculated with 2 × 106 MDA-MB-231 cells and implanted with ATRA-releasing (5 or 10 mg over 21 d) or placebo pellets 3 weeks later (arrow). n = 4. (f,g) Tumor sizes (f, top; measured weekly for 7 weeks), quantitative curves of tumor volume (f, bottom) and representative immunoblots of expression of exogenous and endogenous Pin1 as well as cyclin D1 in xenograft tumors (g) from nude mice inoculated with MDA-MB-231 cells stably expressing Flag-Pin1 or control vector (vec) and, beginning 1 week later, treated with ATRA implants for 7 weeks. n = 3 for f. ***P < 0.001, as determined by Student’s t-test. Error bars, mean ± s.d.
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that contained less Pin1 (Fig. 4f). Similar results were also obtained by treating NB4 human APL cells with ATRA in vitro (Supplementary Fig. 9). Notably, neither Pin1 nor PML–RAR-α was overexpressed in individuals who were in complete APL remission (Fig. 4f–h). Thus, Pin1 inhibition by ATRA, other chemical compounds or shRNA caused PML–RAR-α degradation in APL mouse models and human APL cells in vitro as well as in APL patients.
ATRA as a candidate breast cancer therapyGiven that Pin1 regulates numerous cancer-driving molecules in solid tumors (Supplementary Fig. 1), we hypothesized that ATRA might have anticancer activity against other malignancies. We explored this in breast cancer because of the substantial oncogenic role of Pin1 in this disease9,10,12. We treated nine different human normal and breast cancer cell lines with ATRA and examined cell proliferation by the colorimetric MTT assay. Non-transformed MCF10A and HMLE cells were highly resistant to ATRA, and different malignant cells showed differential susceptibility to ATRA (Fig. 5a).
Compared to MCF10A and HMLE cells, Pin1 was overexpressed in all breast cancer cells11 (Fig. 5b). These cells expressed similar levels of cytochrome P450–dependent retinoic acid-4-hydroxylase (Fig. 5b), and treatment with its inhibitor, liarazole, resulted in generally additive effects with ATRA (Supplementary Fig. 10c), suggesting that differences in ATRA metabolism probably do not account for the observed difference in ATRA sensitivity. Because the Pin1–ATRA co-crystal structure revealed that the carboxyl group of ATRA formed salt bridges with K63 and R69 of Pin1 (Fig. 1i), both of which are responsible for binding the phosphate of Pin1’s pS71 (ref. 7) (Supplementary Figs. 2a and 10a), we examined the possibility that S71 phosphorylation affects ATRA sensitivity. Indeed, the levels of S71 phosphorylation in different cell lines were inversely corre-lated with ATRA sensitivity (Fig. 5b). Given that S71 in Pin1 is phos-phorylated by DAPK1 (ref. 7), a tumor suppressor whose expression is often lost in solid tumors28, we examined the expression of Pin1 and DAPK1 in human TNBC tissues (Supplementary Table 4). High Pin1 but low DAPK1 expression was detected in most breast cancer tissues with an inverse correlation (n = 48) (Fig. 5c and Supplementary Fig. 10b).
To examine whether the inhibitory effects of ATRA on breast cancer cell growth are related to RAR activation, we again used Ro-415253 and AC-93253. As in APL cells, neither compound had obvious effects on the ability of ATRA to induce Pin1 degradation or inhibit prolifera-tion of breast cancer cells (Supplementary Fig. 10d–g).
To further support the thesis that ATRA targets Pin1 in breast cancer, we next examined the effect of ATRA on the abundance of a set of oncogenes and tumor suppressors whose stability is regulated by Pin1 in breast cancer2,3. Indeed, treatment with ATRA caused a dose-dependent decrease in the abundance of Pin1 and its substrate oncoproteins, including cyclin D1 (ref. 12), HER2 (ref. 14), ER-α (ref. 18) AKT16, NF-κB p65 (ref. 13), c-Jun11, and PKM2 (ref. 19), as well as an increase in the abundance of Pin1 substrate tumor suppres-sors such as SMAD2 and SMAD3 (ref. 17) and SMRT15 in all three ATRA-sensitive cancer cell lines (Fig. 5d). ATRA had no appreciable effects on MCF10A cells (Fig. 5d). To further support the notion that these effects are due to Pin1 ablation, we stably introduced Tet-on Pin1 shRNA into these cells. Inducible Pin1 KD had the effects on the proteins encoded by oncogenes and tumor suppres-sors that were similar to those of ATRA treatment (Fig. 5d,e); these effects were rescued by reconstitution of shRNA-resistant Pin1, but not its W34/K63A mutant (Fig. 5f). Thus, ATRA selectively ablates
active nonphosphorylated Pin1 and thereby inhibits multiple cancer- driving pathways in estrogen receptor (ER)-positive, HER2-positive, and triple-negative human breast cancer cells.
To determine whether ATRA inhibits breast tumor growth in vivo, we used the MDA-MB-231 and MDA-MB-468 human TNBC cell lines; we selected TNBC because it has the worst prognosis and the fewest treatment options. In pilot experiments, MDA-MB-231 cells were subcutaneously injected into the flanks of female nude mice, and beginning 1 week later mice were treated with either ATRA (33.0 µmol/kg) or vehicle intraperitoneally three times a week for 8 weeks. ATRA had only modest antitumor activity (Supplementary Fig. 10h), which is consistent with the findings from clinical trials that ATRA has moderate efficacy against advanced breast cancer35,52; this could be due to its short half-life of ~45 min in humans53.
We thus implanted either ATRA-releasing or placebo pellets (to maintain a constant level of the drug) 1 week after injecting TNBC cell lines into the flanks of nude mice. We followed tumor growth for 8 weeks after implantation of ATRA pellets. ATRA pellets potently and dose-dependently inhibited tumor growth and reduced the abun-dance of Pin1 and its substrate cyclin D1 in tumors derived from both MDA-MB-231 (Fig. 6a,b) and MDA-MB-468 cells (Fig. 6c,d), as compared to placebo pellets. Similar dose-dependent inhibition of tumor growth was also observed when ATRA was first administered 3 weeks after MDA-MB-231 tumor cell inoculation, when tumors were already formed (Fig. 6e). To test whether the antitumor activ-ity of ATRA against breast cancer was mediated by Pin1, we stably expressed Pin1 in MDA-MB-231 cells before injecting them into the flanks of nude mice. Pin1 overexpression markedly increased tumor growth (by approximately eightfold), which again was effectively inhibited by ATRA in a dose-dependent manner (Fig. 6f). ATRA also dose-dependently reduced both endogenous and exogenous Pin1 and endogenous cyclin D1 (Fig. 6g). Thus, ATRA has potent anti-tumor activity against TNBC through ablation of Pin1 and its multiple cancer-driving pathways at the same time.
DISCUSSIONThe use of ATRA in APL therapy has been described as the first example of targeted therapy in human cancer36–38, but its drug target(s) remain elusive. Notably, retinoic acid–mediated transactivation is dispensa-ble for leukemia initiated by PML–RAR-α (ref. 54). ATRA’s ability to activate RARs and induce leukemia cell differentiation can be uncoupled from its ability to induce PML–RAR-α degradation, inhibit APL stem cells, and treat APL39,40. ATRA’s ability to activate RARs cannot explain its activity to regulate the protein stability of other oncogenic41,42 and tumor-suppressive43 molecules. Finally, regular ATRA, even with a half-life of 45 min, has moderate but detectable efficacy against solid tumors in some clinical trials, and new genera-tions of supposedly much more potent retinoid derivatives that target RARs or RXRs show little efficacy35,52,55–57.
Our mechanism-based screening has led to the unexpected discovery that ATRA directly binds, inhibits and ultimately degrades active Pin1 selectively. This selectivity was confirmed by solving the Pin1–ATRA co-crystal structure, which reveals that the carboxylic and aromatic moieties of ATRA occupy the Pin1 substrate phosphate- and Pro-binding pockets in the Pin1 active site, respectively, and that S71 phosphorylation prevents ATRA from binding Pin1 by blocking the carboxyl-binding pocket. Notably, the ATRA carboxylic moiety mimics S71 phosphorylation by the tumor suppressor DAPK1, which inhibits Pin1 activity and oncogenic function7. Furthermore, ATRA bound and inhibited Pin1 with Ki and Kd values of 0.5–0.8 µM in vitro.
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An ATRA-releasing formulation effectively treated in situ APL and TNBC mouse models, which reportedly produces a 0.6 µM drug plasma concentration40. ATRA effectively degraded Pin1 and PML–RAR-α in APL cells in human patients at standard doses, which reportedly produce a 1.2 µM drug plasma concentration58. Thus, Pin1 is a critical direct target of ATRA for inducing PML–RAR-α degrada-tion and treating APL, and it is likely to be the long-sought-after drug target of ATRA in the treatment of APL (Supplementary Fig. 11a). Of note, it has been reported that Pin1 inhibition enhances the responses of APL cells to ATRA via stabilization of PML–RAR-α (ref. 59), which is consistent neither with the previous findings that PML–RAR-α causes APL and that ATRA induces PML–RAR-α deg-radation to treat APL39,40 nor with our current findings that ATRA induces PML–RAR-α degradation by directly binding to and degrad-ing Pin1. Moreover, that study did not suggest that ATRA directly targets Pin1 (ref. 59).
Our findings offer a promising new approach for targeting a common oncogenic mechanism to stop numerous cancer-driving molecules and inhibit cancer stem cells at the same time (Supplementary Fig. 11a,b), which is critically needed for treating aggressive or drug-resistant cancers1. Although ATRA has only modest antitumor activity, ATRA-releasing pellets potently inhibit TNBC tumor growth by ablating Pin1 and its targets. Notably, liposomal ATRA, which has a longer half-life, has significant efficacy in APL patients even as a single-agent front-line therapy60. Consistent with this finding is that regular unmodified ATRA has moderate but detectable efficacy against advanced breast cancer in trials35,52. It would be interesting to examine whether cells from those patients who do respond to ATRA treatment show Pin1 degradation, and whether outcomes improve if one could select those patients who respond to ATRA treatment. These results underscore the importance of developing ATRA that has a longer half-life, as doing so would improve its anticancer potency.
Our ATRA–Pin1 co-crystal structure provides insight that ATRA binds to Pin1 by taking advantage of the substrate phosphate- and proline-binding pockets in the Pin1 active site. However, unlike a substrate, the carboxylic and aromatic moieties in ATRA are linked by double bonds, which cannot be isomerized by Pin1. As a result, ATRA may be trapped in the Pin1 active site and inhibit its catalytic activity, ultimately leading to Pin1 degradation, which is supported by the requirement of ATRA binding for ATRA-induced Pin1 degrada-tion. These results could help explain why ATRA seemed to be more potent in vivo than in enzyme-based assays in vitro. Furthermore, the ATRA–Pin1 structure helps explain why the new retinoid derivatives fenretinide and bexarotene33,34 exhibit much lower affinity for Pin1 than ATRA, which may contribute to their failure in the treatment of solid tumors35,56,57.
Our findings also provide a strong rationale for developing more potent and specific Pin1-targeted ATRA derivatives for cancer ther-apy. Comparisons of the Pin1 structures with ATRA and other potent in vitro Pin1 inhibitors27 have identified additional modifications that can be introduced into ATRA to increase its affinity and specificity for Pin1 while reducing its affinity for RARs and possibly improving its half-life. Notably, it is unlikely that ATRA-like Pin1 inhibitors would have major general toxicity because of their selectivity for the active form of Pin1 that is overexpressed in many cancer cells, in addition to the fact that Pin1-KO mice have no obvious defects for an extended period of time25,26. Indeed, ATRA37,38 and even liposomal ATRA60 have not been reported to cause major toxicity.
In summary, we showed that ATRA directly binds, inhibits and ulti-mately degrades the active form of Pin1 that is overexpressed in many
cancer cells to exert potent anticancer activity against APL and TNBC, probably by blocking multiple cancer-driving pathways at once. As regular ATRA has a short half-life of 45 min with moderate anticancer activity against solid tumors in humans, our results provide a rationale for developing either ATRA that has a longer half-life or more potent and specific Pin1-targeted ATRA derivatives for cancer treatment.
METHODSMethods and any associated references are available in the online version of the paper.
Accession codes. Gene Expression Omnibus: Coordinates have been deposited with accession code GSE63059 (array). Protein Data Bank: Pin1, 4TNS.
Note: Any Supplementary Information and Source Data files are available in the online version of the paper.
ACKNoWLEdGMENTSWe thank W.G. Kaelin Jr., N. Gray, J. Clardy and A. Chakraborty for constructive advice; and H. de Thé (INSERM) for RAR-α, RAR-β, and RAR-γ triple-KO MEFs originally generated by P.A. Chambon (Université de Strasbourg); C. Ng for assistance with immunostaining and T. Garvey for editing the manuscript. S.W. is a recipient of a Susan G. Komen for the Cure postdoctoral fellowship (KG111233). The work is supported by grants from the US National Institutes of Health (R01CA167677, R03DA031663 and R01HL111430 to K.P.L.).
AUTHoR CoNTRIBUTIoNSS.W. designed the studies, performed the experiments, interpreted the data, and wrote the manuscript; S.K. helped characterize ATRA binding to and inhibition of Pin1; L.K., J.G. and M.R. helped design and conduct APL-related experiments; W.L. and Y.Z. determined the Pin1–ATRA co-crystal structure; M.N., M.L., Y.Y., A.K., H.H., and C.H.C. provided various technical assistances; M.-H.Y and T.H.L. performed Pin1 and DAPK1 immunostaining; G.B. and H.W. helped analyze Pin1 and ATRA binding; N.J.M. and S.C. provided advice on the FP-HTS screen; E.M.R. and F.L.-C. provided human APL samples; L.C.C. advised the project; P.P.P. advised the project, interpreted the data and reviewed the manuscript; X.Z.Z. developed the original Pin1 FP-HTS and worked with S.W. to identify ATRA; X.Z.Z. and K.P.L. conceived and supervised the project, designed the studies, interpreted the data, and wrote the manuscript.
CoMPETING FINANCIAL INTERESTSThe authors declare competing financial interests: details are available in the online version of the paper.
Reprints and permissions information is available online at http://www.nature.com/reprints/index.html.
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ONLINE METHODSCell culture and reagents. 293T, AU565, BT474, HCC1937, MCF7, MDA-MB-231, MDA-MB-468, SKBR3 and T47D cells (originally obtained from ATCC and maintained by K.P.L) were cultured in DMEM, NB4 cells (obtained from P.P.P.) was cultured in RPMI-1640, and HMLE and MCF10A cells were cultured in F12/DMEM. All mediums were supplemented with 10% FBS and all of the cells were cultured at 37 °C in a humidified incubator contain-ing 5% CO2. HA-Pin1 (Supplementary Figs. 6 and 7) has been previously described7,61. 13cRA, ATRA, EGCG and Juglone were purchased from Sigma. ATRA-releasing pellets were from Innovative Research of America. All muta-tions were generated by site-directed mutagenesis. Antibodies against vari-ous proteins were obtained from the following sources: mouse monoclonal antibodies: Pin1 obtained as previously described12; α-tubulin, β-actin, Flag (M2), γ-tubulin (GTU-88) from Sigma; cyclin D1 (DCS-6), SMRT (1212) from Santa Cruz Biotechnology; Smad (18/Smad2/3) from BD Biosciences; rabbit antibodies: ER-α (Sp1) from Thermo Scientific; HER2 (C-18), PML (H-238), RAR-α (C-20) from Santa Cruz Biotechnology; Akt (9272), c-Jun (9165), PKM2 (3198) from Cell Signaling Technology; NF-κB/p65 (EP2161Y) from Epitomics; Cytochrome p450 (2E1) from Abcam; DAPK1 (DAPK-55) from Sigma. Antibodies against pS71 Pin1 were obtained as previously described7. AC-93253 and Ro-415253 were purchased from Sigma-Aldrich.
Fluorescence polarization–based high-throughput screen. The N-terminal HiLyte Fluor 488–, fluorescein- or TAMRA-labeled peptides (pTide) had a four-residue sequence core structure of Bth-D-phos.Thr-Pip-Nal, which was synthesized using peptide synthesis companies. This sequence was optimized for solubility and binding to Pin1. A selected set of ~8,200 compounds at the Institute for Chemistry and Cell Biology (ICCB-Longwood Screening Facility) at Harvard Medical School was used as the library source. For the screening assay, a solution containing 250 nM glutathione S-transferase (GST)-Pin1 or its PPIase, 5 nM labeled peptide, 10 µg/ml bovine serum albumin, 0.01% Tween-20 and 1 mM DTT in a buffer composed of 10 mM HEPES, 10 mM NaCl and 1% glycerol (pH 7.4) was used. Measurements of FP and FA were made in black 384-well plates (Corning) using the EnVision reader. Compounds were trans-ferred to plates using a custom-built Seiko pin-transfer robot at the ICCB-Longwood Screening Facility. The assay can tolerate up to 10% DMSO. The Z’ is around 0.70 and consistent for day-to-day performance. The coefficient of variation is in the range of 4–5%. Candidates were ranked based on z-score, obtained with the following formula: z-score = (x – µ)/σ, where x is the raw score, µ is the mean of the population, and σ is the s.d. of the population.
Ki values obtained from the FP assay results were derived from the Kenakin Ki equation: Kenakin Ki = (Lb)(EC50)(Kd)/(Lo)(Ro) + Lb(Ro–Lo + Lb–Kd), where Kd [M]: Kd of the probe, EC50 [M]: obtained from FP assay, total tracer Lo [M]: probe concentration in FP, bound tracer Lb [M]: 85% of probe concentration binds to target protein, total receptor Ro [M]: Pin1 concentration in the FP assay, as described30.
Photoaffinity labeling with [3H]ATRA. Photoaffinity labeling of Pin1 with radiolabeled ATRA was performed as described32, with minor modifications, outlined below. 10 pmol of Pin1 was incubated in microcentrifuge tubes with a series of concentrations of all-trans-[11,12-3H]retinoic acid (PerkinElmer, 43.7 Ci/mmol) in 20 µl of the FP assay buffer at 23 °C with agitation for 2 h in the dark. The caps of the microcentrifuge tubes were opened, and the samples were placed on ice and exposed to Electrophoresis System 365/254 nm UV hand lamp (Fisher Scientific) suspended 6 cm above the surface of the liquid for 15 min. The samples were boiled in SDS sample buffer, followed by separation on standard SDS–PAGE gels. The gels were dried and then used for fluorography at −80 °C for 5 d and quantified using Quantity One from Bio-Rad.
PPIase assay. The PPIase activity on GST-Pin1, GST-FKBP12, or GST- cyclophilin in response to 13cRA or ATRA was determined using the chymotrypsin-coupled PPIase activity assay with the substrates Suc-Ala-pSer-Pro-Phe-pNA, Suc-Ala-Glu-Pro-Phe-pNA or Suc-Ala-Ala-Pro-Phe-pNA (50 mM) in buffer containing 35 mM HEPES (pH 7.8), 0.2 mM DTT and 0.1 mg/ml BSA at 10 °C, as described previously5. The Ki value obtained from the PPIase assay was derived from the Cheng–Prusoff equation, Ki = IC50/ (1 + S/Km), where Km
is the Michaelis constant for the used substrate, S is the initial concentration of the substrate in the assay, and IC50 is the half-minimal inhibitory concentration of the inhibitor, as described62.
Inhibition of cell proliferation. Cells were seeded with a density of 3,000 cells per well in 96-well flat-bottomed plates and incubated for 24 h in 10% FBS– supplemented DMEM culture medium. Cells were then treated with ATRA alone or in combination with other drugs. Control cells received DMSO at a concentration equal to that in drug-treated cells. After 72 h, either the number of cells was counted after trypsin digestion or medium containing 0.5 mg/ml 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyl-2H-tetrazolium bromide was added to each well for 2 h incubation at 37 °C, followed by removing the media before adding 200 µl DMSO. Absorbance was determined at 570 nm.
Immunoprecipitation and immunoblotting. Cells were polyethylenimine (PEI)- or lipofemamine-transfected with 8 µg of various plasmids and incu-bated in 10 cm dishes for 24 h, followed by drug treatment as needed. When harvested, cells were lysed for 30 min at 4 °C in IP lysis buffer (50 mM HEPES, pH 7.4, 150 mM NaCl, 1% Triton X-100 and 10% glycerol) with freshly added phosphatase and protease inhibitors consisting of 100 µM 4-(2-aminoethyl)-benzenesulfonyl fluoride, 80 nM aprotinin, 5 µM bestatin, 1.5 µM E-64 protease inhibitor, 2 µM leupeptin, 1 µM pepstatin A, 2 mM imidazole, 1 mM sodium fluoride, 1 mM sodium molybdate, 1 mM sodium orthovanadate, and 4 mM sodium tartrate dihydrate (all from Sigma). After centrifugation at 13,000g for 10 min, one-tenth of the supernatant was stored as input, and the remainder was incubated for 12 h with M2 Flag agarose (Sigma). After brief centrifugation, immunoprecipitates were collected, extensively washed twice with the afore-mentioned lysis buffer, suspended in 2× SDS sample buffer (100 mM Tris-HCl, pH 6.8, 4% SDS, 5% β-mercaptoethanol, 20% glycerol, and 0.1% bromophe-nol blue), boiled for 10 min, and subjected to immunoblotting analysis. Equal amounts of protein were resolved in 15% SDS-polyacrylamide gels. After electrophoresis, gels were transferred to nitrocellulose membranes using a semi-dry transfer cell (Bio-Rad). The transblotted membrane was washed twice with Tris-buffered saline containing 0.1% Tween-20 (TBST). After blocking with TBST containing 5% bovine serum albumin (BSA) for 1 h, the membrane was incubated with the appropriate primary antibody (diluted 1:1,000) in 2% BSA–containing TBST at 4 °C overnight. After incubation with the primary antibody, the membrane was washed three times with TBST for a total time of 30 min, followed by incubation with horseradish peroxidase (HRP)-conjugated goat anti-rabbit or anti-mouse IgG (diluted 1:2,500) for 1 h at room temperature. After three extensive washes with TBST for a total time of 30 min, the immu-noblots were visualized by enhanced chemiluminescence.
Immunostaining and fluorescence microscopy. Human APL samples were kindly provided by E.M.R. Tissue samples were washed with PBS and fixed with 4% paraformaldehyde at room temperature for 20 min, followed by per-meabilization and blocking with PBS containing 0.1% Triton X-100 and 5% FBS for 1 h. After another wash with PBS, immunostaining was performed by incubating the cells with mouse anti-Pin1 (Homemade, 1:1,000), or rabbit anti-PML (H-238, Santa Cruz; 1:100) primary antibodies at 4 °C overnight. Primary antibodies were diluted in PBS containing 0.1% Triton X-100, 0.2% BSA, 0.5 mM PMSF and 1 mM dithiothreitol. After washing with PBS, secondary Alexa Fluor 488–conjugated goat anti-mouse antibodies or Alexa Fluor 564–conjugated goat anti-rabbit antibodies (Invitrogen; 1:200) were added at room temperature for 2 h. Samples were nuclear counterstained with 4,6-diamidino-2-phenylindole (DAPI), mounted and visualized with LSM510 confocal imaging system. For centrosome duplication assays, NIH 3T3 cells were used as described previously7. Briefly, cells were synchronized in G1/S phase by adding 10 µg/ml aphidicolin for 24 h, then fixed with 4% paraformaldehyde at room temperature for 20 min, stained for centrosomes with anti-γ-tubulin antibodies (GTU-88, Sigma; 1:100) and analyzed by confocal microscopy.
Immunohistochemistry. Tissue microarrays with a cohort of 48 human TNBC tissues were purchased from US Biomax. Immunohistochemical staining for Pin1 was performed as described previously23,63. In brief, deparaffinization and hydration were carried out with xylene, 100%, 75% and 50% ethanol and water,
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sequentially. Antigen retrieval was performed by boiling samples in an auto-clave device for 20 min in 1× antigen retrieval Citra (Biogene). Samples were blocked with PBS containing 5% goat serum and 0.1% Triton X-100, followed by incubation with anti-Pin1 antibody (against nonphosphorylated Pin1, 1:200) or anti-DAPK antibody (1:500) at 4 °C in a humidified chamber for 12 h. After extensive washes with PBS, samples were incubated with biotinylated secondary antibody for 2 h and visualized with the Vectastain ABC kit and DAB-staining solution (Vector Laboratories). In each sample, expression of Pin1 or DAPK1 was semi-quantified manually in a double-blind manner as high, medium or low according to the standards presented in Figure 6c. The correlation of Pin1 expression and DAPK1 expression in 48 human TNBC tissues was analyzed by Spearman’s rank correlation test (P < 0.001).
Genome-wide gene expression profiling. NB4 human APL cells were treated with 10 µM ATRA (Sigma-Aldrich) or doxycycline-induced Pin1 knockdown for 3 d, and total RNA was extracted with the Trizol reagent according to the manufacturer’s instructions. The samples were then processed using Affymetrix GeneChip WT PLUS Reagent Kit, followed by Hybridization Wash and Stain Kit. Microarray expression profiles were collected using Affymetrix Human Transcriptome Array 2.0. Original CEL files were analyzed by the Affymetrix software programs Expression Console and Transcriptome Analysis Console. Microarray data have been deposited in NCBI Gene Expression Omnibus with series accession number GSE63059. Genes with lower expression in Pin1-KD or ARTA-treated cells than in vec or DMSO-treated cells with a fold change < 0.5 (P < 0.05) were selected as downregulated ones, and those with higher expression in Pin1 KD or ARTA-treated cells than in vec or DMSO-treated cells with a fold change > 2 (P < 0.05) were selected as upregulated ones.
Crystallization and complex structure determination for Pin1 PPIase domain with ATRA. Pin1 PPIase domain (residue 51–163) was cloned into a pET28a derivative vector with an N-terminal hexahistidine tag followed by recognition sequences by thrombin and PreScission 3C proteases and then followed by the recombinant gene. Mutations of K77Q, K82Q were created by QuikChange site-directed mutagenesis (Agilent Technologies).
The PPIase K77/82Q was purified similarly to a previous published method29 with minor modifications. Briefly, PPIase K77/82Q was overexpressed in E. coli BL21 (DE3) cells with 1 mM isopropyl-β-D-thiogalactopyranoside (IPTG) induction at 16 °C overnight. Cell lysate was first purified using nickel affinity chromatography. The elution was a dialysis in a buffer of 20 mM HEPES, 100 mM NaCl, 8 mM β-Mercaptoethanol (pH 8), while protein was treated with PreScission Protease (GE) overnight at 4 °C. After His-tag removal, Pin1 PPIase K77/82Q was separated from untruncated protein by a second round of nickel affinity chromatography, and then purified by size exclusion chroma-tography columns Superdex 75 (GE Healthcare).
Purified PPIase K77/82Q was concentrated to 15 mg/ml. ATRA dissolved in DMSO at the concentration of 1mM and was mixed with protein solution and incubated on ice for 3 h before setting up trays. Incubated protein was co-crystalized by vapor diffusion using a hanging drop of 1 µl protein-ATRA plus 1 µl well solution. The complex formed crystals in solutions containing 0.2 M ammonium sulfate, 0.1 M HEPES (pH 7–8.5) and 0.9–1.4 M sodium citrate after micro-seeding using apo-PPIase domain crystals. The crystals were cryoprotected by the addition of 30% glycerol in mother liquor and vitrified in liquid nitrogen before data collection.
X-ray diffraction was collected from synchrotron radiation at beamlines 5.0.2 of the Advanced Light Source (Berkeley, California) with 3 × 3 CCD array detectors (ADSC Q315R). Data were processed and scaled using the HKL 2000 software suite64. Data collection statistics are summarized in Supplementary Table 1.
The structure of PPIase K77/82Q bound with ATRA was determined by molecular replacement with PPIase K77/82Q (PDB: 3IKG) as the search model using program Phaser from the CCP4 package suite65. The structure was refined with the Refmac5 program from CCP4 package and iterative model building in COOT66,67. The final structure was evaluated by both PROCHECK68 and MolProbity69. Refinement statistics are summarized in Supplementary Table 1. The Pin1–ATRA structure was deposited into the Worldwide Protein Data Bank with the PDB code of 4TNS.
Animal studies. For xenograft experiments, 2 × 106 of MDA-MB-231 or MDA-MB-468 parent cells or MDA-MB-231 cells expressing Pin1 or control vectors were injected subcutaneously into flank of 8-week-old female BALB/c nude mice (Jackson Laboratories). One week later, tumor growth was just about notable by sight, and mice were randomly selected to receive ATRA or control treatment. For intraperitoneal injection, vehicle or 12.5 mg/kg ATRA was administered three times a week for 8 weeks. For implantation, placebo, 5 or 10 mg over 21 d of ATRA-releasing pellets (Innovative Research of America) were implanted subcutaneously in the back of nude mice. Tumor sizes were recorded weekly by a caliper for up to 8 weeks and tumor volumes were calculated using the formula: L × W2 × 0.52, where L and W represent length and width, respectively. For NB4 cell transplantation, 8-week-old NOD.Cg-prkdcscidll2rgtm1Wjl/SzJ (termed NSG) mice (Jackson Laboratory)70 were used as recipients after sublethal irradiation at 350 Gy. Each mouse was transplanted with 5 × 105 NB4 cells stably express-ing Tet-on shPin1 via retro-orbital injection. Five days later, when transplanted cells had established, mice were randomly selected to receive regular or doxy-cycline-containing food and survival curves were recorded. For PML–RAR-α transgenic cell transplantation, C57BL/6 mice were given 350 Gy irradiation followed by transplantation with 1 × 106 APL cells from CTSG-PML-RARA transgenic mice50,71. After 5 d, when transplanted cells had established, mice were randomly selected to receive placebo (21 d placebo-releasing pellets, sub-cutaneously), ATRA (5 mg of 21d ATRA-releasing pellets, subcutaneously), EGCG (12.5 mg/kg/d, intraperitoneally), or Juglone (1 mg/kg/d, intravenously). Mice were sacrificed after 3 weeks, when APL blast cells appeared in a peripheral blood smear of placebo-treated mice. Spleen weight was measured and bone marrow was collected for immunoblotting of PML–RAR-α and Pin1 expression. No animals were excluded during the experiments. Animal work was carried out in compliance with the ethical regulations approved by the Animal Care Committee, Beth Israel Deaconess Medical Center, Boston, USA.
Human APL samples. Bone marrow aspirates were obtained with informed consent from the iliac crests of individuals in whom the diagnosis of acute pro-myelocytic leukemia was suspected based on a morphological evaluation of their peripheral blood smears. Immediately after the procedure, therapy with ATRA was started72,73. Second bone marrow aspirate samples were obtained on day 3 or 10 of ATRA therapy. Samples tested positive for the PML–RAR-α rearrangement by RT-PCR. The human sample collection was approved by the Institutional Review Board at the University of São Paulo (HCRP no. 13496/2005) or at Tor Vergata University (IRB no. 12/07).
Statistical analysis. Experiments were routinely repeated at least three times, and the repeat number was increased according to effect size or sample variation. We estimated the sample size considering the variation and mean of the samples. No statistical method was used to predetermine sample size. No animals or samples were excluded from any analysis. Animals were randomly assigned to groups for in vivo studies; no formal randomization method was applied when assigning animals for treatment. Group allocation and outcome assessment was not done in a blinded manner, including for animal studies. All data are presented as the means ± s.d., followed by determining significant differences using the two-tailed student’s t-test or analysis of variance (ANOVA) test, where *P < 0.05, **P < 0.01, ***P < 0.001.
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